Phylogenomic Analyses Reveal the Evolutionary Originof the Inhibin a-Subunit, a Unique TGFb SuperfamilyAntagonistJie Zhu1, Edward L. Braun2, Satomi Kohno2, Monica Antenos1, Eugene Y. Xu1, Robert W. Cook1, S. Jack
Lin3, Brandon C. Moore2, Louis J. Guillette, Jr.2, Theodore S. Jardetzky3, Teresa K. Woodruff1*
1 Department of Obstetrics and Gynecology, Feinberg School of Medicine, Northwestern University, Chicago, Illinois, United States of America, 2 Department of Biology,
University of Florida, Gainesville, Florida, United States of America, 3 Department of Structural Biology, Stanford University School of Medicine, Stanford, California, United
States of America
Abstract
Transforming growth factor-beta (TGFb) homologues form a diverse superfamily that arose early in animal evolution andcontrol cellular function through membrane-spanning, conserved serine-threonine kinases (RII and RI receptors). Activin andinhibin are related dimers within the TGFb superfamily that share a common b-subunit. The evolution of the inhibin a-subunit created the only antagonist within the TGFb superfamily and the only member known to act as an endocrinehormone. This hormone introduced a new level of complexity and control to vertebrate reproductive function. The novelfunctions of the inhibin a-subunit appear to reflect specific insertion-deletion changes within the inhibin b-subunit thatoccurred during evolution. Using phylogenomic analysis, we correlated specific insertions with the acquisition of distinctfunctions that underlie the phenotypic complexity of vertebrate reproductive processes. This phylogenomic approachpresents a new way of understanding the structure-function relationships between inhibin, activin, and the larger TGFbsuperfamily.
Citation: Zhu J, Braun EL, Kohno S, Antenos M, Xu EY, et al. (2010) Phylogenomic Analyses Reveal the Evolutionary Origin of the Inhibin a-Subunit, a Unique TGFbSuperfamily Antagonist. PLoS ONE 5(3): e9457. doi:10.1371/journal.pone.0009457
Editor: Sudhansu Kumar Dey, Cincinnati Children’s Research Foundation, United States of America
Received December 4, 2009; Accepted February 1, 2010; Published March 4, 2010
Copyright: � 2010 Zhu et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricteduse, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: This work was supported by grant numbers R01HD37096 and R21ES014053 from the National Institutes of Health. The content is solely theresponsibility of the authors and does not necessarily represent the official views of the National Institutes of Health. The funders had no role in study design, datacollection and analysis, decision to publish or preparation of the manuscript.
Competing Interests: The authors have declared that no competing interests exist.
* E-mail: [email protected]
Introduction
The common ancestor of the TGFb superfamily arose early in
animal evolution, before bilaterians and cnidarians diverged [1]. The
TGFb gene family is large and has members that encode dimeric
ligands of membrane-spanning serine-threonine kinases (RII and RI
receptors) that regulate diverse cellular functions at the autocrine or
paracrine level [2,3]. The 42 members of the TGFb superfamily
include the TGFbs, bone morphogenic proteins (BMPs), growth and
differentiation factors (GDFs), activins, inhibins, anti-Mullerian
hormone (AMH), Nodal, and the leftys [4,5]. TGFb superfamily
members are synthesized as pre-prohormones that are extensively
processed to a ‘mature’ C-terminal active form (Fig. S1). Each
processed monomer has six to nine conserved cysteine residues in the
mature domain that form a cysteine knot motif, which is
characteristic of all TGFb superfamily members (reviewed in [6])
and is crucial to both the three-dimensional structure of the monomer
and formation of the active dimer. Activin and inhibin are members
of the TGFb superfamily linked by structure, function and phylogeny.
Inhibin and its unique a-subunit adhere to many but not all of the
rules established for the other superfamily ligands. Following the
stepwise evolution of the inhibin a-subunit provides a unique vantage
point in understanding the constraints on the other members of this
extraordinarily powerful and highly conserved family of ligands.
The activin subgroup of the TGFb superfamily comprises four
polypeptide subunits in mammals: bA, bB, bC, and bE [7,8] are
112–116 amino acids in length and contain nine conserved
cysteines. Activins are homo- and heterodimers of these subunits
(activin A and activin B are the dominant ligands of the
reproductive axis). Activins are made locally and constitutively
and then bind cell surface activin type II receptors (ActRIIA/B)
and subsequently stimulate activin type I receptors (ActRIA/IB) to
initiate intracellular signaling events [3,9]. Inhibin is the only
TGFb superfamily ligand assembled from two relatively dissimilar
subunits; it shares a b-subunit with activin but couples with a
dissimilar a-subunit [10]. In mammals, the inhibins (inhibin A
[a-bA] and inhibin B [a-bB]) also bind the activin type II receptors
(ActRIIA/B) in addition to an accessory binding protein called
betaglycan, but do not activate downstream signaling proteins
[11,12,13]. In this manner, inhibin acts as an antagonist that
blocks the ability of activin to bind and activate its receptors.
The dominant role of activin is to locally and constitutively
stimulate follicle-stimulating hormone (FSH) synthesis and secre-
tion from pituitary gonadotrope cells [14,15]. The pituitary
hormone FSH is released in a cyclic manner (monthly in humans
and every 4 days in the mouse) and activates a new set of growing
ovarian follicles. In direct response to FSH, the follicles produce
inhibins that are secreted into the peripheral circulation and block
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activin-regulated receptors on the gonadotrope. In this manner,
peripheral inhibin blocks activin-dependent FSH production at
precise times during the reproductive cycle in a classical negative
feedback loop [16,17,18,19]. Loss of activin or inhibin is
catastrophic to the reproductive cycle of all mammals [20].
The activin bA2 and bB2subunits show a high degree of
sequence conservation in all invertebrates and vertebrates that
have been examined [21]. In contrast, the a-subunit of inhibin
does not appear in lower invertebrates and mammalian ortholo-
gues are quite different from the b-subunits except in the spacing
of the cysteines, which has lead to speculation regarding the origin
of the a-subunit and the evolution of its functional characteristics.
For example, the mature avian a-subunit is 113 amino acids in
length (similar to the b-subunits) but contains seven cysteines (the
mature b-subunit contains nine cysteines) [22]. Moreover, all b-
subunits have a helical ‘wrist region’ necessary for receptor
binding [23]. This wrist region is the location of an intriguing
difference between a- and b-subunits that is unexpected from an
evolutionary perspective. This wrist region is missing from the a-
subunits of non-mammalian vertebrates whereas a precise
insertion that effectively replaces the wrist region has occurred at
this position in mammalian a-subunits [22]. However, the
mammalian insertion is a proline-rich sequence that is not
homologous to the wrist region helix of b-subunits. Moreover,
both avian and mammalian mature a-subunit proteins have an
extended N-terminus and antibodies directed at this region
neutralize the biological activity of the hormone [24,25]. These
structural domains provide clues to the function of the ligands and
a roadmap of evolutionarily flexible domains.
The goal of this phylogenomic analysis of the inhibin/activin
genes was to establish the timing of gene duplications, as well as
the number and types of changes that occurred after duplication.
If the inhibin a-subunit arose from a b-subunit ancestor around
the origin of the vertebrates, what were the steps that converted
the activin b-subunit into an a-subunit? Sequence changes that
occurred after the origin of the inhibin a-subunit should represent
changes associated with the acquisition of novel functions of this
subunit. Is it possible to account for the unique functions of the
inhibin a-subunit not present in the activin b-subunit or any other
TGFb ligand at the sequence level? Specific functions unique to
the inhibin a-subunit are the ability to heterodimerize exclusively
with activin b-subunits (forming a-b dimers) but not homodimer-
ize (a-a) [26] and to act as an antagonist of activin. Can we assign
these functions through an analysis of the structural domains that
were iteratively assigned to the a-subunit during animal evolution?
These questions can be addressed by phylogenomic analysis and
the construction of a set of inhibin ligands based on the results of
our phylogenomic analysis to rationally forward and reverse
engineer molecules that mimic evolutionary intermediates of the
a-subunit. Combining a structure-function analysis with phyloge-
nomics provides a powerful way to understand the contributions of
iterative evolutionary changes that contributed to the development
of complex reproductive strategies in vertebrates.
Results
Regions of Interest and Models of EvolutionaryModification
A number of plausible hypotheses for the evolution of vertebrate
inhibin/activin genes are possible (Fig. 1), most of which postulate
the origin of the critical a-subunit occurred after duplication of a
gene encoding b-subunit-like TGFb homolog. Indeed, the
genomic location of the human INHA gene (chromosome 2q),
which encodes the inhibin a-subunit, suggests a specific version of
this ‘‘b-subunit duplication’’ model since INHA is located in a
region with extensive intragenomic homology to chromosomes 7,
12q, and 17q centered on paralogous Hox gene clusters [27]. The
origin of these paralogous Hox clusters has been suggested to
reflect two whole genome duplications (WGDs) early in vertebrate
evolution [28,29,30] (Fig. 1A). Human b-subunit genes are located
near three of these Hox clusters on chromosomes 2q (INHBB,
encoding bB), 7p (INHBA, encoding bA), and 12q (the adjacent
INHBC and INHBE genes, encoding bC and bE) (Fig. 1B). This
‘‘early vertebrate b-subunit duplication’’ model of inhibin/activin
evolution suggests neo-functionalization (the origin of a novel
function; [31] immediately after duplication. The hypothesis that
the vertebrate genome arose by WGD is not universally accepted
[32,33], although a variant of the model of inhibin/activin
evolution involving the duplication of genomic blocks rather than
WGD remains possible. Alternatives to the early vertebrate b-
subunit duplication model, involving either an earlier origin within
the deuterostomes (Fig. 1C) or even an origin before the
divergence of protostomes and deuterostomes (Fig. 1D), are also
possible. In fact, one such alternative is suggested by the existence
of a clade of a-subunit and insect Dawdle homologs (also called
‘‘activin-like proteins’’) in a phylogenetic analysis of TGFb
Figure 1. Plausible models of evolution for the inhibin/activingene family. A) Simplified species tree for animals based upon recentlarge-scale analyses [93,94,95]. ‘‘WGD’’ indicates the position of thewhole genome duplications [29] uniting vertebrates and letters indicatealternative models for the origin of the a-subunit (the letters usedcorrespond to the parts of this figure). B) Cladogram showing theexpected inhibin/activin phylogeny given the ‘‘early vertebrateduplication’’ model. Two rounds of WGD have been suggested tocharacterize vertebrates [28,29,30] and we show the only topologyconsistent both with two rounds of WGD and a clade containing INHBA-INHBB (the latter clade is strongly-supported in this study and in otherstudies; [27,96]. C) Cladogram showing the expected phylogeny ofinhibin/activin genes given the ‘‘deuterostome duplication’’ model,which places the a-subunit origin before the early vertebrate WGDs.Several versions of this model are possible (i.e., the a-subunit origincould predate the divergence of vertebrates from urochordates,cephalochordates, or even echinoderms). D) Cladogram showing theexpected inhibin/activin phylogeny given the ‘‘early animal duplication’’model, which places the a-subunit origin before the divergence ofdeuterosomes and protostomes. The ‘‘Dawdle orthologue’’ hypothesisis a version of this model. Additional duplications may have occurred inany of these models. Biased estimation of the gene tree or samplingvariance may cause estimates of the gene tree to deviate from any ofthese idealized model trees.doi:10.1371/journal.pone.0009457.g001
Evolution of Inhibin a-Subunit
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homologues [34]. This ‘‘Dawdle orthologue’’ hypothesis can be
viewed as a specific version of the ‘‘early animal b-subunit
duplication’’ model (Fig. 1D). Either of the alternative models
suggests a relatively early duplication that established b-subunit
paralogues that subsequently underwent changes and acquired
specific functions to become an a-subunit. The major open
question regarding the evolution of the vertebrate inhibin/activin
gene family are when the duplication that led to the a-subunit
occurred and what the implications of that timing were for
functional changes that led to the origin of inhibin.
Phylogenomics Reveals the Origin of the Inhibina-Subunit
To distinguish among alternative models for the origin of the
inhibin a-subunit, we identified genes encoding inhibin/activin
homologues in annotated genome sequences and aligned the TGFbdomains encoded by those genes using conserved structural elements,
like the conserved cysteine residues and the W-X-X-W element, as
landmarks (data not shown). The large number TGFb homologues
and conflict among previous estimates of TGFb phylogeny
homologues [34,35,36] suggest the evolution of the TGFb super-
family has been complex. To better understand the evolution of this
family, we extracted a large set of TGFb homologues from available
animal genome sequences and identified those that had a human
inhibin/activin gene as a top hit (this corresponds to the bidirectional
BLAST best hit criterion) (Fig. S2A, Table S1). These database
searches revealed a number of potential inhibin/activin genes in
several different invertebrate groups (Figs. 2A and 2B). However, all
putative inhibin/activin genes appeared to have the structural
features typical of b-subunits and genes that clearly encode a-
subunits could only be detected in vertebrates using BLAST (Figs. 2B
and S2A). A phylogenomic analysis using the maximum-likelihood
[37] criterion suggested that one of the lancelet (Branchiostoma floridae)
inhibin/activin genes is likely to represent an a-subunit orthologue
(Figs. 2A, 2B, S2B and S2C), a hypothesis further corroborated by the
presence of a specific insertion (Fig. S2B). Insertion and deletion
(indel) events are typically considered very reliable evolutionary
markers [38], so this insertion provides strong evidence for a specific
relationship between the lancelet protein and vertebrate a-subunits.
Unlike vertebrate a-subunits, which have seven cysteine residues, the
putative lancelet a-subunit orthologue has nine cysteines, suggesting it
has retained an ancestral condition of the inhibin/activin family that
is typical of b-subunits (Fig. S2B). The lancelet a-subunit does have an
N-terminal extension, although it is rich in charged residues, unlike
the proline-rich N-terminal extension of vertebrate a-subunits
(Figs. 2A and S2C). The lancelet a-subunit also has a partial deletion
of the wrist helix (Fig. 2A). Since we were unable to identify inhibin/
activin genes in other deuterostome invertebrates, like tunicates or sea
urchins (Fig. 2B), there must have been multiple instances of gene loss.
This is not unexpected since gene loss has played a major role in
eukaryotic evolution [39] and one of the lineages highlighted by this
study (tunicates) is known to have undergone substantial gene loss
[40]. Indeed, the distribution of genes encoding both a- and b-
subunits (Fig. 2B) suggests the evolution of the TGFb superfamily
reflects a complex birth-death process with many instances of gene
duplication and gene loss.
Our phylogeny is consistent with an origin of the inhibin a-
subunit prior to the divergence of vertebrates and cephalochor-
dates (Fig. 2B), and the duplication leading to the vertebrate a-
and b-subunit families could be even more ancient (Figs. 1C and
1D). As expected based upon the short sequences being used [41],
phylogenetic analyses of the TGFb superfamily have limited
power and we were unable to exclude alternative models with a
more ancient origin of the a-subunit lineage (Fig. S2A).
Regardless, the duplication that led to the a-subunit is likely to
predate the putative WGDs (Fig. 1A) that resulted in the
duplications of the regions surrounding the Hox gene clusters,
thought to have occurred close to the origin of vertebrates [42,43].
Since the only clear invertebrate a-subunit is the lancelet
sequence, the ‘‘deuterostome b-subunit duplication’’ model
(Fig. 1C), which postulates an origin of the a-subunit by
duplication of a b-subunit gene within deuterostomes, probably
represents the simplest hypothesis for the origin of the gene
encoding the inhibin a-subunit.
The Inhibin a-Subunit Accumulated Major Insertion andDeletion Changes after Duplication
Comparison of a-subunit and b-subunit sequences revealed a
surprisingly limited set of sites where the sequences overlap in a
clear manner, with many regions aligning very poorly (Figs. 2A
and S2D). The deletion event near the critically important wrist
region and the complex indels near the extended N-terminal
region are of interest from both evolutionary and functional
perspectives (Figs. S2C and S2D). Here, we compared the inhibin
a- and bA-subunits to determine what evolutionary changes in the
a-subunit resulted in the substantial expansion of functions among
the inhibins and activins within the TGFb superfamily [6].
Proline-rich regions near the N-terminal region of the mature a-
subunit and the wrist region were particularly striking, especially
when we examined a large number of inhibin a-subunit sequences
from non-mammalian vertebrates to clarify the evolution of these
regions (Figs. 2A and S2D, Table S2). The phylogenomic
distribution of these proline-rich regions suggests that the N-
terminal extension arose relatively early in the evolution of the a-
subunit whereas the wrist region arose much later (Figs. 2C and
S3). This alignment suggests that indel changes represent a major
type of change that differentiate the inhibin a-subunit and b-
subunits, and the distribution of the indels suggest they could have
played an important role in changing a-subunit function.
The evolutionary histories of the a-subunit and b-subunits differ
fundamentally, since there are four mammalian genes encoding b-
subunits (INHBA, INHBB, INHBC and INHBE) and a single gene
encoding the a-subunit (INHA). The ML estimate of phylogeny for
the mature inhibin/activin protein is consistent with an origin of
three major b-subunit clades (corresponding to the mammalian bA,
bB, and bC/bE groups) early in vertebrate evolution (Fig. 1B),
probably during the WGD [29] (Fig. 1A). The bC and bE groups
reflect a subsequent gene duplication event that occurred after
mammals’ divergence from other vertebrate groups (Fig. S2A). In
sharp contrast, we identified a single a-subunit in all taxa we
examined (Fig. S2) and the phylogeny included many expected
groups (major clades like mammals and teleosts were present;
Figs. 2C and S3). Although there were areas of incongruence with
the accepted vertebrate phylogeny (e.g., the deepest-branching
amniotes were birds and a squamate-mammal clade was found)
bootstrap support for these apparent conflicts were limited (Figs. 2C
and S3). There are two potential explanations for conflict between a
gene tree and the expected species tree. First, the conflict could be
genuine and reflect processes that lead to incongruence between
gene trees and species trees [44]. Second, the conflict could be only
apparent, reflecting an inaccurate estimate of the gene tree either
due to bias or limited power. We expected phylogenomic analyses of
the inhibin/activin family to have limited power given the limited
alignment length [41], and we could not reject the likely species tree
based upon bootstrap support (Figs. 2C and S3). These results
suggest there have been multiple gene duplications during the
evolution of the b-subunit but few duplications and losses during the
evolution of the a-subunit.
Evolution of Inhibin a-Subunit
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Figure 2. Phylogenomic analyses revealed a complex series of indels that correlated with major events in the evolution of theinhibin a-subunit. A) Alignment of mature inhibin/activin proteins showing expanded sampling within the vertebrates for the a-subunit. Thealignments were optimized based upon the highly conserved cysteine residues [97] and the highly conserved W-X-X-W motif and R-X-X-Rproconvertase enzyme cleavage site (bold). N-terminal region and wrist region were highlighted with grey shade. B) Cladogram showingevolutionary relationships among animals with annotated genome sequences available using a topology based upon a consensus of recent analyses[93,94,95]. The two major clades of bilaterian animals (deuterostomes and protostomes) are highlighted. Numbers of proteins that exhibit clearhomology to inhibin/activin queries in BLASTP searches are shown to the right, with the number of those proteins that have a human inhibin/activina- or b-subunit as their top hit when used as a query in BLASTP searches of human proteins indicated in parentheses. Thus, numbers in parenthesesreflect the number of proteins that are candidates for inhibin/activin a- or b-subunits using a bidirectional BLAST criterion. The likely origins of a- andb-subunits based upon the phylogenomic analyses reported here are indicated using the relevant Greek letters, and the timing of the whole genomeduplications uniting vertebrates are indicated as ‘‘WGD’’. The branch at the base of the tree is hatched because the position of the root is unclear[93,94,95]. C) Schematic of the ML estimates the phylogeny for inhibin/activin proteins, emphasizing the occurrence of indels in the mature proteinregion during the evolution of the gene family. Support for specific groups is indicated as the percentage of 100 bootstrap replicates, with onlyvalues $50% indicated. The starlet sea anemone (Nematostella) activin homolog is the sister of the lancelet (Branchiostoma) b-like protein and isindicated using a light line since it is probably placed incorrectly (note that bootstrap support is limited). The dashed arrow indicates the likelyposition of the sea anemone activin; this position is more likely because it minimizes the number of gene duplications and losses given the likelyorganismal phylogeny shown in B). A detailed version of this phylogeny is provided in Fig. S3.doi:10.1371/journal.pone.0009457.g002
Evolution of Inhibin a-Subunit
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Mechanisms of a-Subunit EvolutionThe most striking differences between vertebrate a- and b-
subunits are the loss of two cysteine residues and the presence of
several large indels (Figs. 2A and 2C). These changes occurred at
distinct times in the evolutionary history of the a-subunit. The
lancelet a-subunit, the only likely inhibin a-subunit we could
identify in an invertebrate, does have an N-terminal extension
despite having nine cysteine residues similar to the b-subunit.
However, it is unclear whether the lancelet N-terminal extension is
homologous to the vertebrate N-terminal extension, since the
lancelet sequence is rich in charged (especially acidic) residues
whereas the vertebrate insertion is proline-rich (Fig. S2C). The
second major insertion in the a-subunit, which corresponds to
amino acids 68–85 of the human protein (Figs. 2A and 2C), is
called the wrist region because it falls between the cysteines where
the wrist helix in the inhibin b-subunit is located [45,46]. Non-
mammalian inhibin a-subunits (i.e., those from fish, amphibians,
reptiles, and birds) lack the wrist region insertion but some type of
wrist region insertion is present in all known mammalian inhibin
a-subunits (Figs. 2A, 2C and S2D).
Multi-residue insertions like those observed in the a-subunit are
thought to be rare genomic changes [38]. Indeed, both of the
major indel events can be mapped as single events on the inhibin
a-subunit phylogeny (Figs. 2C and S3) and the fact that marsupials
and placental mammals share the long wrist region insertion
provides further support for the now well-accepted therian clade
[47,48] that excludes prototherians (contra initial mitogenomic
analyses, which supported a marsupial-prototherian clade; [49].
However, the evolution of the proline-rich wrist region (PWR)
appears to have been highly dynamic. There is a relatively short (5
amino acids) insertion in the platypus but longer (8–19 amino
acids) insertions are present in therians. The shortest therian
insertion (8 amino acids in the Tammar wallaby, Macropus eugeni)
probably reflects a secondary deletion given the existence of a
longer (14 amino acid) insertion in other marsupials and the
phylogenomic position of the wallaby (Figs. 2A and 2C).
Emphasizing the dynamic nature of this region, the monotreme
and therian wrist-region insertions could have been independent
or the shorter platypus insertion could reflect a deletion. However,
a stepwise model involving multiple insertions is both simple and
consistent with the data. The N-terminal insertion is even more
complex, since distinctive insertions are present in the lancelet and
in vertebrates suggesting multiple origins of an insertion in this
region (Figs. 2A, 2C and S2C). Like the wrist region insertion,
there appears to have been a number of indels in this region. This
includes independent insertions in teleosts and mammals as well as
deletions in the proline-rich N-terminal insertion (Fig. 2A).
Examination of the inhibin/activin alignment in an explicit
phylogenomic framework emphasizes that there have been
multiple indels concentrated in two specific regions and that some
of the largest insertions in these regions are correlated with specific
events (i.e., the a-subunit origin at the base of the vertebrates and
the origin of therian mammals (Figs. 2C and S3).
The existence of multiple indels in specific regions of proteins
raises questions about their role in the evolution of a-subunit
function and the basis for their recurrence in the same regions.
Although multi-residue indels are viewed as rare genomic changes
that can be especially valuable for defining clades in phylogenomic
trees, multiple insertions can occur in the same position in proteins
and these insertions are evident even in studies that have used
insertions as phylogenetic markers [50,51]. Protein structural
features are likely to constrain insertions to specific parts of
proteins, creating the appearance of recurrent indels. Likewise,
insertions are likely to share certain features that may make
convergent insertions difficult to identify. These common features
might include large numbers of hydrophilic residues (because
insertions tend to form exposed loops) or residues that can form
flexible linkers like proline and glycine. Indeed, the wrist region
insertion is both proline-rich and less conserved than the
remainder of the mature a-subunit as one might expect if the
function of the wrist region is to act as a flexible linker (flexible
sequences often evolve at high rates [52]). The N-terminal
insertion is more conserved than the wrist region insertion, but
it is also proline-rich. Mechanisms of indel mutations remain
poorly understood, although a positive correlation between GC-
content and coding region indel rate has been documented [53].
Mammalian INHA third codon positions are GC-rich, as are the
two major insertions. Indeed, amino acids over-represented in
flexible regions (i.e., glycine and proline) have GC-rich codons,
suggesting that insertions in GC-rich coding regions could have a
higher likelihood of producing a flexible linker that can be
tolerated by proteins. Indeed, it is reasonable to postulate a
feedback loop in which an insertion creates a flexible linker and a
region prone to further indel changes. Regardless of larger-scale
differences among genes in indel probability, the base composition
of major INHA insertions is consistent with the composition of the
gene (i.e., the wrist-region insertions are 93% GC [14 of 15 nt,
including 5 of 5 third positions] in the platypus and 75% GC [36
of 48 nt, including 10 of 16 third positions] in the human). Like
any novel mutation, indels could be fixed by drift, but any changes
to a flexible linker that alters the activity of the gene product to
produce an advantageous variant would be fixed by natural
selection. Thus, regardless of the mechanism of most indel
mutations, there are clear reasons to examine the impact of
specific indels upon the function of the inhibin a-subunit.
Taken as a whole, our phylogenomic analyses emphasize that the
inhibin a-subunit lineage arose by an ancient b-subunit duplication,
probably early in deuterostome evolution. Our analyses also indicate
that there have been multiple duplications of genes encoding the b-
subunit and few (if any) duplications of a-subunit encoding genes.
Some of the most striking changes during a-subunit evolution have
been large insertions, making it reasonable to postulate that the
differences in biological activity between the a-subunit and the activin
b-subunits and other TGFb ligands reflect these insertions.
Forward-Engineered bA-Subunits with a-Subunit IndelsDetermine Homodimerization Potential
To test the hypothesis suggested by our phylogenomic
analyses—that insertions played a major role in the origin of a-
subunit function after the a-subunit arose by duplication of
another TGFb superfamily member (b-subunit gene)—we con-
structed chimeric inhibin/activin genes and examined their
biological activity. We modeled the iterative evolution of the
inhibin a-subunit using the human bA-subunit as a template.
Human bA-subunits were used because they exhibit more than
90% sequence identity to other vertebrates in the mature region
[21,54], are well-characterized functionally [55,56,57] and appear
to have undergone limited change since the common ancestor of
the inhibin/activin family (Fig. 2C). Forward-engineered bA-
subunit mutants were constructed to change the bA-subunit into
an ‘‘a-like’’ subunit. Specifically, we swapped the human a-subunit
N-terminal extension (ext) to the bA-subunit (bAext+/bAext+
mutant), deleted the bA-subunit wrist helix (WH) region
(bAWHD/bAWHD mutant) and swapped the a-subunit proline-rich
wrist region (PWR) into the wrist helix region of the bA-subunit
(bAPWR+/bAPWR+ mutant) (Figs. 3A and S4, Table S3). The bA-
subunit wrist helix region deletion mutant activin (bAWHD/
bAWHD mutant) retained its ability to homodimerize but some
Evolution of Inhibin a-Subunit
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monomer bAWHD product was present (Fig. 3A). The addition of
the a-subunit N-terminal extension region to the bA-subunit also
partially blocked homodimer assembly; however, the addition of
the mammalian a-subunit proline-rich wrist region into the bA
helix region completely blocked dimer assembly (Fig. 3A). These
data are consistent with a model in which early vertebrate inhibin
a-like subunits, which have the wrist helix region deletion but lack
the proline-rich wrist region insertion, gained some potential for
heterodimerization. Addition of the full N-terminal extension then
further reduced homodimerization, while facilitating inhibin a-
subunit heterodimerization with the b-subunit. With the proline-
rich wrist region insertion in the mammalian a-subunit, the a-
subunit was then only able to form heterodimers.
Does Avian Inhibin a-Subunit Retain the Ability toHomodimerize?
Since homodimerization was strongly suppressed when the a-
subunit proline-rich wrist region was swapped into the human bA-
subunit, we asked whether wild-type avian a-subunit (which lacks 4
amino acids in the N-terminal insertion and has no proline-rich wrist
region insertion) was able to homodimerize (Fig. S2D). We cloned the
avian (chicken aChwt) and mammalian (human aHwt) inhibin a-
Figure 3. Cloning and expression of engineered inhibin and activin mutants and their wild type molecules. Schematic representationsof wild type and mutant inhibin/activin molecules along with immunoblots demonstrating their presence in conditioned media from stablytransfected cells. The construct transfected is marked above each lane and the product bands are indicated on the side with lower case letters ornumbers or Greece letters that correspond to the schematic representation on the left. N-linked glycosylation site is denoted by the y symbol. Detailson the design and nomenclature of all the mutants described herein are provided in Fig. S4 and Table S3. A) Left, schematic representations ofdimeric and monomeric forms of wild type activin (bA/bA), activin chimera mutants (bAext+/bAext+ or bAPWR+/bAPWR+) and activin deletion mutant(bAWHD/bAWHD). Right, immunoblot of wild type activin and its chimera and deletion mutants using anti-bA-subunit polyclonal antibody under non-reducing (NR) and reducing (R) conditions. B) Left, schematic representations of wild type chicken inhibin (aChwt/bA) as well as dimeric andmonomeric forms of chicken free a-subunit (aChwt/aChwt, aChwt and Pro-aChwt). Middle, immunoblot of wild type chicken free a-subunit and chickeninhibin using anti-human-a-subunit PO23 monoclonal antibody under non-reducing (NR) or reducing (R) conditions. Far Right, non-reducing SDS-PAGE autoradiograph showing the immunoprecipitation (using anti-a-subunit monoclonal antibody PO23) of [35S]-cysteine-labeled chicken free a-subunit and chicken inhibin A. Controls for the immunoprecipitation experiment are shown in Fig. S5A. C) Left, schematic representations of humanwild type inhibin A (aHwt/bA), chicken wild type inhibin (aChwt/bA), and three deletion mutants of the human a-subunit (aHext2/bA, aHPRW2/bA andaHext-PRW2/bA). Right, immunoblot of wild type human and chicken inhibin A and the human a-subunit deletion mutants using anti-a-subunit PO23monoclonal antibody under non-reducing (NR) condition. Subsequent immunoblots detected with either anti-a-subunit monoclonal antibody PO23or anti-bA-subunit polyclonal antibody under reducing conditions are shown in Fig. S6.doi:10.1371/journal.pone.0009457.g003
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subunits and expressed each species alone or in a bicistronic vector
containing the conserved human bA-subunit (aChwt/bA or aHwt/bA)
(Figs. 3B and 3C). As expected, a-a homodimers were not detected in a
cell line expressing only the mammalian a-subunit [35S] labeling of
cellular proteins (Fig. S5A). Moreover, the heterodimer was the
dominant ligand expressed by the isogenic cell line expressing the avian
a-subunit and bA-subunit cassette (Figs. 3B and 3C). If the avian
(chicken) a-subunit alone was expressed, low, but detectable, aChwt/
aChwt mature homodimers were detected both by immunoblot and
autoradiography following [35S] labeling of cellular proteins (Fig. 3B
and S5A), and were confirmed by MS-MS (Figs. S5B and S5C). These
data are consistent with the notion that the ancestral form of the
inhibin a-subunit retained some ability to homodimerize, although the
dominant secreted form was a heterodimer [58]. Both the N-terminal
insertion and the proline-rich wrist region insertion evolved iteratively
(shorter insertions are present outside of the eutherians; Figs. 2A and
2C) and the homodimerization potential of the a-subunit appears to
have been progressively reduced as the insertions lengthened.
To determine whether the N-terminal extension or proline-rich
wrist region insertions present in the human inhibin a-subunit affect
a-b heterodimer formation, the relevant mutants were cloned into a
bicistronic vector containing the bA-subunit. The mutants tested
were those relevant to our evolutionary model, including a deletion
of N-terminal insertion (aHext2/bA mutant, where amino acids 6–
27 have been deleted), a deletion of the proline-rich wrist region
insertion (aHPWR2/bA mutant, where amino acids 68–85 have
been deleted) or in tandem deletion (aHext-PWR2/bA mutant) (Fig.
S4, Table S3). There were no significant differences between the
inhibin a-subunit deletion mutants and wild-type human inhibin in
terms of heterodimer secretion (Figs. 3C and S6).
Forward-Engineered bA-Subunit Indels Regulate AgonistFunction
We next asked whether the forward-engineered activin
molecules (dimers of bA-subunits containing a-subunit segments)
acted as agonists or antagonists. We tested the same three mutant
classes created by altering the bA-subunit in a manner consistent
with the major evolutionary changes identified in the phyloge-
nomic analysis. Again, these changes were an addition of the N-
terminal insertion (bAext+/bAext+ mutant), deletion of the wrist
helix region (bAWHD/bAWHD mutant) and addition of the proline-
rich a-subunit wrist region insertion (bAPWR+/bAPWR+ mutant)
(Fig. S4, Table S3). Each mutant was tested in a functional assay
that measured FSHb-luciferase activity in a stably transfected
pituitary gonadotrope cell line (2338 FSHb-Luc) [59]. Addition of
the N-terminal insertion reduced the bioactivity of the mutant
activin compared to wild type, but still acted as an agonist (mutant
EC50 = 3.82 nM; wild type EC50 = 1.21 nM) (Table 1). Activity of
the mutant bAext+/bAext+ dimer was not altered with the addition
of the co-receptor betaglycan (EC50 = 5.13 nM) (Table 1). In sharp
contrast, elimination of the wrist helix region of the activin b-
subunits (bAWHD/bAWHD mutant) also eliminated activin agonist
function (Table 2) [23,46]; indeed, this ligand showed weak
antagonist properties (IC50 = 56.7 vs. wild type inhibin A
IC50 = 1.01 nM) (Table 1). We measured the affinities of wild
type activin A, chimera mutant bAext+/bAext+ and deletion mutant
bAWHD/bAWHD for ActRIIB directly using a competition binding
analysis, which revealed a weaker binding constant for the N-
terminal insertion mutant and wrist helix deletion mutant (bAext+/
bAext+ IC50 = 5.99 nM; bAWHD/bAWHD IC50 = 47.6 nM; human
activin A IC50 = 4.14 nM) (Figs. 4A and S7A). Thus, the critical
switch of the a-subunit from an agonist to an antagonist appears to
have coincided with the deletion of the wrist helix region; when
examined in isolation, the N-terminal insertion only changed the
relative strength of bA-subunit agonist activity.
As expected based upon our model of inhibin a-subunit
evolution, placement of the proline-rich wrist region of eutherian
mammals into the bA-helix region (bAPWR+/bAPWR+ mutant)
does not restore the agonist function of the bAWHD/bAWHD
mutant (Table 2). However, bAPWR+/bAPWR+ mutants also did
not form productive homodimers (Fig. 3A). Taken as a whole,
these data indicate that deletion of the wrist helix region was a
critical step for functionally switching the b-subunit from an
agonist to antagonist. Based upon our phylogenomic analyses of
the inhibin/activin gene family, we postulated that the a-subunit
arose by neo-functionalization after duplication a b-subunit gene;
Table 1. Functional assay of wild type and mutant inhibin and activin constructs as measured by FSHb promoter activity in stablytransfected LbT2 cells.
Inhibin A/Activin A wild type andmutants n IC50 (nM) EC50 (nM) F-test p
2BG +BG 2BG +BG
aHwt/bA antagonist 12 1.0160.27a 0.2460.03 - - ,0.05
aChwt/bA strong antagonist 12 0.5760.25ab 0.3660.33 - - .0.05
aHext2/bA weak antagonist 12 4.4662.26c 0.6360.23 - - ,0.05
aHPWR2/bA strong antagonist 12 0.3160.16b 0.2760.14 - - .0.05
aHext-PWR2/bA not an antagonist 12 - - - -
bA/bA agonist 9 - - 1.2160.16A
bAWHD/bAWHD weak antagonist 9 56.7617.9d - - -
bAext+/bAext+ weak agonist 9 - - 3.8260.33B 5.1362.23 .0.05
bAPWR+/bAPWR+ poor dimerization - - - - -
IC50: concentration of ligand causing 50% of the maximum inhibition.EC50: concentration of an agonist producing 50% of the maximum possible response.a,b,c,d represents differences between the groups, p,0.05 by F-test.A,B represents differences between the groups, p,0.05 by F-test.BG = betaglycan, H = human, Ch = chicken, wt = wild type.ext = N-terminal extension, PWR = Proline-rich wrist region, WHD = wrist helix deletion.doi:10.1371/journal.pone.0009457.t001
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these analyses indicate the mutation most likely to be responsible
for this change is the deletion of the wrist helix region.
Reverse-Engineered a-Subunit Indels Identify FunctionalChanges
Because non-mammalian inhibin a-subunits (including those
from fish, amphibians, reptiles and birds) contain the wrist helix
deletion typical of all non-mammalian inhibin a-subunits but
lack the proline-rich wrist region insertion found in mammals
(Figs. 2A and 2C), we wanted to examine the activity of a non-
mammalian a-subunit. The chicken inhibin (aChwt/bA) was
chosen as a representative non-mammalian a-subunit to test in
the FSHb functional assay (since the activin type II receptor and
betaglycan ZP domain are conserved between the avian and
mammal species) for comparison to human inhibin A (aHwt/bA).
We observed two notable differences between the functional
capacity of human inhibin and chicken inhibin. First, chicken
inhibin was a slightly more potent antagonist than human inhibin
(chicken inhibin A IC50 = 0.57 nM; human inhibin A
IC50 = 1.01 nM) (Table 1). Second, while the co-receptor
betaglycan enhanced human inhibin activity (IC50 = 0.24 nM)
as expected [11,60], addition of betaglycan did not further affect
the already high chicken inhibin antagonist function
(IC50 = 0.36 nM) (Table 1).
Interaction of human and chicken inhibins with the ActRIIB
activin receptor was then examined by co-immunoprecipitation.
Whereas chicken inhibin interacted with ActRIIB similarly in the
presence and absence of betaglycan, human inhibin binding to
ActRIIB was enhanced by betaglycan (Figs. 4B and S8). We
measured the affinities of chicken and human inhibin for
ActRIIB directly using a competition binding analysis, which
revealed a stronger binding constant for chicken inhibin
(IC50 = 5.55 nM vs. human inhibin A IC50 = 9.01 nM) (Figs. 4C
and S7B). Thus, the ancestral a-subunit could have been a
slightly stronger antagonist with a higher affinity to the ActRIIB
that was able to act in a betaglycan-independent manner to block
activin action.
Because none of the deletion mutants exhibited problems with
heterodimer formation, we had the opportunity to further test the
impact of specific mutations on inhibin function. We first tested
deletion of the proline-rich wrist region insertion in the human a-
subunit (aHPWR2/bA mutant), which revealed that this mutation
creates a remarkably potent antagonist that no longer requires
betaglycan (IC50 = 0.27 nM with betaglycan vs. IC50 = 0.31 nM
without betaglycan) (Table 1). Moreover, this mutant behaved
similar to chicken inhibin in co-immunoprecipitation assays with
ActRIIB and betaglycan (i.e., there was little difference in
aHPWR2/bA mutant inhibin binding to ActRIIB with or without
betaglycan; Figs. 4B and S8). ActRIIB receptor interaction was at
a very high affinity (aHPWR2/bA IC50 = 2.96 nM vs. wild type
human inhibin A IC50 = 9.01 nM) (Figs. 4C and S7B), consistent
with strong antagonistic function in the FSH bioassay.
Deletion of the N-terminal insertion (aHext2/bA mutant)
resulted in an inhibin that was less bioactive than wild type
inhibin (IC50 = 4.46 nM) (Table 1). This result was consistent with
previous findings that an antibody to the N-terminal extension
region bioneutralized inhibin function [24,25]. The N-terminal
deletion mutant showed increased antagonist function with the
addition of betaglycan, similar to wild type human inhibin
(IC50 = 0.63 nM with betaglycan) (Table 1). The N-terminal
deletion mutant bound to ActRIIB in the co-immunoprecipitation
assay, which increased with the addition of betaglycan (Figs. 4B
and S8). The competition binding affinity of the N-terminal
deletion mutant to ActRIIB was weaker than that observed for
human wild type inhibin A (IC50 = 127.9 nM) (Figs. 4C and S7B).
Elimination of the both the N-terminal insertion and the proline-
rich wrist region insertion (aHext-PWR2/bA mutant) resulted in the
almost complete abolition of inhibin bioactivity, and no activin A-
like agonist function. Betaglycan did not rescue the antagonist
function of this double mutant (Table 2). The double deletion
mutant inhibin A exhibited no binding to ActRIIB, with or
without betaglycan, similar to the empty vector (Figs. 4B and S8).
This mutant did not show any competition binding affinity to
ActRIIB with biotinylated human wild type inhibin A (data not
shown).
Taken as a whole, these experiments emphasize that the
proline-rich insertions found in the inhibin a-subunit of eutherian
mammals resulted in profound changes to the biological activity of
that gene. Specifically, loss of the wrist region helix from the b-
subunit early in the vertebrate evolution, probably prior to the
vertebrate WGD but after the divergence of vertebrates and
cephalochordates, resulted in a shift from an agonist to an
antagonist. The insertion of a long proline-rich sequence into the
area, where the wrist region helix had been located, abolished any
remaining homodimerization potential and allowed interaction
with the betaglycan co-receptor. The insertion in the N-terminal
Table 2. Summary of wild type and engineered inhibin/activin indel mutant.
Mature dimerprocessing
Activin-likefunction Inhibin-like function
Affected bybetaglycan
aHwt/bA Yes No Yes Yes
aChwt/bA Yes No Yes, but stronger than human wild type inhibinA No
aHext2/bA Yes No Yes, but weaker than human wild type inhibinA Yes
aHPWR2/bA Yes No Yes, but stronger than human wild type inhibinA No
aHext-PWR2/bA Yes No No No
bA/bA Yes Yes No No
bAWHD/bAWHD Yes No Yes, but weaker than human wild type inhibinA No
bAext+/bAext+ Yes Yes, but less thanwild type activinA
No No
bAPWR+/bAPWR+ Very little No ND ND
ND = not determined.doi:10.1371/journal.pone.0009457.t002
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region also had an impact on homodimerization capacity and the
activity of the dimer, but functional changes related to insertions in
this region appear to have been more limited in the vertebrate
inhibin a-subunit. Nonetheless, these experiments strongly confirm
the predictions of the phylogenomic study that indels would prove
to be central to functional changes among members of the
inhibin/activin family.
Discussion
Inhibin and the Progressive Evolution of VertebrateReproductive Strategies
Inhibin is the only known endocrine hormone in the TGFbsuperfamily, and based on the present phylogenomic analysis,
likely arose early in vertebrate evolution, before the WGD that
Figure 4. Equilibrium binding of activin and inhibin mutants to ActRIIB using competitive ELISA and immunoprecipitation studies.A) Conditioned media from bA/bA, bAext+/bAext+, and bAWHD/bAWHD expressing cells were quantified and used to compete with biotinylated activinA for binding to ActRIIB. The IC50 for the bA/bA, bAext+/bAext+ and bAWHD/bAWHD competition binding curves were 4.14631.61 nM, 5.9960.98 nM,47.6614.3 nM, respectively, representing a significant difference between all groups as determined by sigmoidal dose-response (variable slope)curve, followed by an F-test (p,0.05). The standard curve for biotinylated activin A binding to the ActRIIB (EC50 = 1.1 nM) is shown in Fig. S7A. B)Top, densitometric analysis of immunoblots (bottom) from three independent experiments, Density was normalized to COS7 cells transfected withempty vector (pcDNA3) and treated with human wild type inhibin A. Asterisks represent statistically significant differences using unpaired t-test(p,0.05). Bottom, the immunoblots of the immunoprecipitation that use monoclonal anti-HA pulldown for lysates from COS-7 cells transfected withActRIIB-HA and either full-length betaglycan or empty vector (pcDNA3). Prior to cell lysis, the cells were treated with inhibin deletion mutants, wildtype human inhibin A, chicken inhibin A or activin A culture media for 2 hours. The western-blots are detected by anti-bA-subunit polyclonalantibody. Controls for the immunoprecipitation studies are shown in Fig. S8. C) Conditioned media from aHwt/bA, aCHwt/bA, aHext2/bA and aHwr2/bA expressing cells were quantified and used to compete with biotinylated inhibin A for binding to ActRIIB. The IC50 for the aHwt/bA, aCHwt/bA, aHwr2/bA and aHext2/bA competition binding curves were 9.0163.72 nM, 5.5561.85 nM, 2.9661.23 nM and 127.9617.80 nM, respectively, representing asignificant difference between all groups as determined by sigmoidal dose-response (variable slope) curve, followed by F-test (P,0.05). The standardcurve for biotinylated inhibin A binding to the ActRIIB (EC50 = 2.73 nM) is shown in Fig. S7B. The graphs shown in (A) and (C) represent the results ofmore than three independent experiments.doi:10.1371/journal.pone.0009457.g004
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characterizes the vertebrate genome. Inhibin plays an essential
role in the negative feedback between the gonads and the pituitary
gland that are necessary for the regulation of reproductive function
[17,61]. FSH in the pituitary is stimulated by locally produced
activin, and ovarian-derived inhibin antagonizes activin in the
pituitary [14,15,17]. Interestingly, the inhibin a-subunit and the b-
subunit of FSH might have arisen at similar times; both molecules
arose by gene duplications early in vertebrate evolution [62], and
are necessary for the integration of information from distal tissues
to control reproduction.
To learn more about inhibin and provide clues about its origins
and iterative changes that resulted in the creation of a powerful
endocrine antagonist within the TGFb family and the reproduc-
tive axis, we used phylogenomics in combination with rationally
designed mutants that were tested in well-characterized functional
and binding assays (Figs. 5A and 5B). We learned two important
pieces of information from this work. First, the b-subunit-like
ancestor of the inhibin a-subunit underwent at least two key
changes that resulted in the acquisition of novel functions (neo-
functionalization) and led to the emergence of the inhibin a-
subunit (Fig. 5A). Second, a hierarchy of functions can be assigned
to specific molecular signatures (the indels) that allow us to
understand the changes that occurred during the evolution of this
molecule (Fig. 5C).
Our data are consistent with a model in which the invertebrate
b-subunit underwent a series of changes to become a functional a-
subunit. The lancelet a-subunit-like sequence, united with
vertebrate a-subunit and, based upon the results of phylogenomic
analyses (Figs. 2C and S3), has a partial deletion of the helix and a
distinct N-terminal extension, suggesting it may be a ‘‘missing
link’’ between the a- and b-subunits. Loss of the helix region
present in the b-subunits, which is responsible for receptor
Figure 5. Schematic diagrams illustrating the proposed model for the evolution of the inhibin a-subunit. A) Indels that have had amajor impact upon a-subunit function are indicated on the deuterostome tree. Two major indel changes (the N-terminal insertion and wrist helixregion deletion [1] and the proline-rich wrist region insertion [2]) occurred at distinct times in evolution. B) Schematic representation of the evolutionof inhibin a-subunit. The graph includes the forward engineered mutants used in our experiment that are representative of a-like transient forms.Non-mammalian a-subunit structure is represented by the chicken a orthologue, and mammalian a-subunit structure is diagramed by the human aorthologue. C) Evolution from inhibin b-subunit to inhibin a-subunit involves the loss of the ability to homodimerize. Concurrent with this evolutionwas a change in the bioactivity from agonist to strong antagonist that is betaglycan independent, and later to weak antagonist that requiresbetaglycan for maximal antagonistic function. The indel in the evolution of inhibin a-subunit strongly affects inhibin function.doi:10.1371/journal.pone.0009457.g005
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binding, particularly to the activin type I receptor (ALK4)
[10,45,46], switched the function of the molecule from an agonist
to an antagonist and facilitated heterodimer formation. Loss of the
helix region alone, however, was not sufficient to achieve full a-
subunit functionality; the N-terminal extension developed in
parallel to strengthen the antagonistic properties of the a-subunit
and further favor heterodimer formation (Fig. 5).
The inhibin a-subunit in non-mammalian vertebrates has a
deletion of the wrist region (e.g., the chicken a-subunit) and
inhibin dimers containing these a-subunits do not require the
accessory protein betaglycan. As the wrist region became
populated by a series of proline amino acids, likely to have first
appeared as short insertions in the common ancestor of
monotreme-therians and later expanded to form larger insertions
in metatherian and eutherian mammals, the inhibin molecule was
able to better coordinate betaglycan receptor binding (Fig. 5C).
We predict that most non-mammalian vertebrates exhibit a more
powerful betaglycan-independent antagonism similar to that
exhibited by avian inhibin. By contrast, mammalian inhibin is a
relatively weaker antagonist of activin signaling that is dependent
upon betaglycan as a co-receptor (Fig. 5C). Inhibin has a role
limited to the time of sexual maturity in birds and progesterone
regulates pituitary function after that time. Indeed, immunoneu-
tralization of chicken inhibin brings the animals into egg-laying
earlier [63] and the birds continue to cycle after entering sexual
maturity. Thus, a strong endocrine antagonist is necessary for the
onset of normal sexual maturity in the chicken, but is not necessary
in the normal adult cycle [64,65,66]. Conversely, immunoneu-
tralization of mammalian inhibin causes a prompt rise in FSH and
the end of adult cyclicity [25,67]. Thus, a weaker antagonist may
be necessary to ensure an on-and-off rate necessary for a repetitive
cycle that must be reset rapidly (up to every four days in some
rodents) [68].
A fundamental question in evolutionary biology is whether
innovations (or novelties) can be accommodated within the neo-
Darwinian framework [69,70]. There has been vigorous debate
regarding the roles of drift and selection in the evolution of genes,
genomes and organisms [71,72], but the mechanisms by which
genomic changes result in evolutionary novelties remain unclear.
The evolution of inhibin, a TGFb ligand that can act as an
endocrine hormone, represents a potentially important molecular
innovation. The molecular basis of this innovation appears to be
gene duplication followed by a sequence change that inhibits
homodimerization (Fig. 5C), a process that conforms to one of the
well-established ‘‘principles governing molecular evolution’’ [73].
The origin of novel functions after gene duplication is easily
accommodated within modern evolutionary theory [74,75,76].
This is true even for some of the more recently suggested pathways
for the origin and preservation of duplicate genes [74,75,77,78].
Ultimately, changes to a duplicate gene that appear to conform to
expected types of change [73] led to the advent of the inhibin a-
subunit which resulted in an innovation that allowed communi-
cation between gonads and pituitary, itself a novelty at the level of
the whole organism. This communication between the gonads and
pituitary represents the evolution of an important endocrine
feedback mechanism, perhaps the first negative feedback loop to
emerge in vertebrate animals.
Here we have presented evidence regarding the nature of
specific evolutionary changes that led to the origin of the inhibin a-
subunit in the vertebrate lineage. These molecular changes appear
to result in innovations, both in molecular function and in the
specialization of the reproductive axis, specifically the ability to act
as an endocrine antagonist. The subsequent molecular changes
leading to a ‘‘mammalian’’ mode of inhibin signaling also
emphasized the role of co-option or recruitment in genetic
innovation. Specifically, the wrist region insertion resulted in the
recruitment of betaglycan, which had an established role in TGFbsignaling [60,79,80], to enhance the inhibin signaling system. Our
experiments indicate that well-established processes such as gene
duplication followed by sequence changes can explain these
evolutionary novelties. However, a striking difference between the
most prominent sequence changes noted in this study of inhibin
evolution and those noted in many other studies is the importance
of indels rather than amino acid substitutions [10]. In contrast to
the excellent methods available to examine positive selection for
amino acid sequences [81,82] that have been successfully applied
at a genomic scale [82,83] tools that can be used to examine the
potential functional impact of indels remain poorly developed.
Despite the relative paucity of analytical tools, it is clear that indels
make an important contribution to genomic divergence [84] and
to changes in protein function [83,85,86]. Our results are striking
in that they directly implicate a relatively poorly characterized
type of sequence change in the origin of inhibin function.
Nonetheless, the fact that the impact of indels on protein function
remains poorly characterized relative to the impact of amino acid
substitutions does not a change the fact that indels can be
accommodated within the neo-Darwinian paradigm.
Development of the inhibin a-subunit resulted in the first
negative feedback peptide-hormone based endocrine system in
animals by providing a circulating, gonadally-derived antagonist to
the pituitary-made activin. Neo-functionalization of the inhibin a-
subunit expanded the possibilities for animal reproductive
strategies. The early a-subunits, characterized by N-terminal
insertion and the wrist region deletion were able to block activin
receptors without an accessory protein, although they retained
some ability to homodimerize. The wrist region underwent
additional insertions in the mammals that led to further functional
modification, including the inability to homodimerize and a
requirement for the betaglycan co-receptor, further attesting to the
importance of this feedback loop for animal reproductive
strategies. Combining phylogenomic analysis together with
functional assays of a rationally designed set of mutant proteins
provided a remarkable opportunity to understand the origins of
the evolutionary complexity of the inhibin system and provide
information about the reproductive evolutionary changes needed
to create this agonist-antagonist pair.
Materials and Methods
Gene Sequencing and Phylogenomic AnalysisA diverse set of TGFb homologues were retrieved from
complete genome sequences (downloaded from NCBI [http://
www.ncbi.nlm.nih.gov] and the DOE Joint Genome Institute
[http://www.jgi.doe.gov]) using BLASTP searches [87]. Initial
sequence alignments were obtained using MAFFT [88,89] and
optimized by eyes using the conserved cysteine residues, the
conserved W-X-X-W motif and conversed RXXR proconvertase
enzyme cleavage site. To examine the phylogenomic distribution
of specific a-subunit indels, additional sequences of the mature
inhibin a-subunit were determined by RT-PCR or PCR of
genomic DNA. It was reasonable to use genomic DNA as a
template because the a-subunit comprises two exons in all species
studied to date and the splice sites are found upstream of the
region encoding the N-terminus of the mature region. The ML
estimate of phylogeny was obtained using PhyML 3.0 [89] using
the JTT model [90] of amino acid evolution with the rates at
different sites drawn from a C-distribution, which was judged to be
the best-fitting model using standard criteria [41]. Support for
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specific groups was assessed using 100 bootstrap replicates and
indels were mapped onto the phylogeny using the maximum
parsimony criterion [91,92].
Mutagenesis and Protein ProductionActivin bA-subunit chimeras containing regions of the inhibin a-
subunit were produced by amplifying appropriate regions of the
inhibin a-subunit (the N-terminal insertion and the proline-rich
wrist region insertion) using primers containing partial activin bA-
subunit and partial inhibin a-subunit sequences. The QuikChange
procedure (Stratagene, La Jolla, California, USA) was then followed
using the amplicons as the primers in a PCR reaction in which a
pcDNA3 plasmid containing the bA gene was the template. Inhibin
deletion mutants were also made with the QuikChange method
using primers spanning, but not including, the region of the a-
subunit that was removed (Table S3). The template for the reaction
was a pcDNA5/FRT bicistronic plasmid containing the human a-
and bA-subunits separated by an internal ribosomal entry site [37].
The full-length chicken a-subunit cDNA was kindly provided by Dr.
Patricia A. Johnson (Cornell University, Ithaca, New York, USA)
and subcloned into the first multiple cloning site of the inhibin A
bicistronic plasmid replacing the human a-subunit cDNA. Either
full-length human or chicken a-subunit was subcloned into
pcDNA5/FRT. Mutations were verified by sequencing in an
ABI3100 Capillary DNA Sequencer. CHO cells were transfected
with the activin chimera mutants using Lipofectamine 2000
(Invitrogen, Carlsbad, California, USA). Transfected CHO cells
were selected using G418 antibiotic (500 mg/ml) and carried in
DMEM-F12 media supplemented with 5% fetal bovine serum, 1%
penicillin/streptomycin. CHO Flp cells were transfected with
inhibin deletion mutant plasmids or inhibin free a-subunit plasmids
with pOG44 plasmid and were selected using hygromycin B
antibiotic (500 mg/ml) and carried in F12 media supplemented with
10% FBS and 1% penicillin/streptomycin. Three isogenic cell lines
were generated for human inhibin A, chicken inhibin A, human
inhibin free a-subunit, chicken inhibin free a-subunit and each
inhibin deletion mutant [37]. At confluence, media was exchanged
for DMEM-F12 or F-12 serum-free media and grown for 3–4 days
before collection. Following collection, serum-free media was
dialyzed into 50 mM Tris, 150 mM NaCl, and the protein solution
was concentrated by Amicon Ultra Centrifugal Filter Units
(Millipore, Billerica, MA, USA) [10]. The concentrated media
was directly used in the bioassay and binding assay.
Immunoblot Analysis of Ligands and ProteinConcentration Determination
Wild type, chimera mutant, and human a-subunit deletion
mutant ligands were electrophoresed on 4–12% SDS-PAGE gels
under non-reducing conditions and transferred to nitrocellulose
for immunoblot analysis. The ligands were detected using a
polyclonal antibody to the inhibin bA-subunit antibody (kindly
provided by Dr. W. Vale, The Salk Institute, La Jolla, California,
USA) followed by horseradish peroxidase-conjugated anti-rabbit
secondary antibody (Zymed, San Francisco, California, USA). The
secondary antibody was detected using ECL (Pierce Chemical Co.,
Rockford, Illinois, USA) and exposed at varying time points to X-
ray film (Kodak, Rochester, New York, USA). Densitometric
analysis of signals was quantified using the KODAK 4000 MM
Digital Imaging System and analyzed using Kodak Imaging
Software (version 4.0.1 Kodak, Rochester, NY) [37]. Purified
human wild type activin A and inhibin A of known concentration
(from our laboratory) was used as a standard to quantify the
amount of ligand from wild type activin A or inhibin A and
mutants culture media concentrated and dialyzed under non-
reducing conditions. The inhibin A deletion mutant and activin
chimera mutants were detected using one of three antibodies to
the mature a-subunit: anti-a-subunit R1 monoclonal antibody
(from Serotec, Raleigh, North Carolina, USA) directed toward the
extended region, anti-a-subunit PO14 monoclonal antibody
(provided by Dr. David Phillips, Monash Institute, Australia)
directed toward the wrist region, and anti-a-subunit PO23
monoclonal antibody (provided by Dr. David Phillips, Monash
Institute, Australia) directed toward the C-terminal region (Fig.
S1).
Metabolic In Vivo Labeling and SubunitImmunoprecipitation
Isogenic cultures of cells expressing wild type human inhibin A,
chicken inhibin A, human inhibin free a-subunit and chicken
inhibin free a-subunit in 12 well plates were continuously cultured
in the presence of [35S]-cysteine. After 48 hours, media was
collected and immunoprecipitated with anti-a-subunit monoclonal
antibody PO23 or normal mouse IgG, and then carried with the
Protein G Sepharose 4 Fast Flow (GE Healthcare, Sweden). Beads
were then boiled for 5 min, centrifuged and the supernatants were
analyzed on 4–12% SDS-PAGE gel under non-reducing condi-
tions. Autoradiography using X-ray film (Kodak, Rochester, New
York, USA) was conducted for about 7 days.
Tandem MS-MS Fragment Mass MappingCulture media from cells stably expressing the chicken free a-
subunit was collected and immunoprecipitated with an anti-a-
subunit monoclonal antibody PO23 and subjected to SDS-PAGE
under non-reducing conditions. After the gel was stained by
Coomassie Blue (Bio-Rad, Hercules, California, USA), the 30 kDa
band was excised and sent to the protein and nucleic acid facility
at the Stanford University Medical Center (Stanford, California,
USA). The band was digested with trypsin and subjected to
tandem MS-MS analysis.
Luciferase Assays of FSHb ExpressionThe bioactivity of wild type and chimera proteins was
determined using LbT2 gonadotrope cell line stably transfected
with the 2338 region of the FSHb promoter conjugated to a
luciferase reporter [59]. Cells were cultured as reported previously
[59], and treated for 6 hours with various concentrations of wild
type activin A or media from cells expressing the activin A chimera
mutants, or activin A (0.4 nM) with various concentrations of
media from cells expressing wild type human inhibin A, chicken
inhibin A, or human inhibin A deletion mutants at 37uC in serum-
free DMEM/F12 (1:1), phenol red-free (Invitrogen, Carlsbad,
California, USA), supplemented with 1% penicillin/streptomycin.
Following treatment, cells were lysed in GME buffer with 1%
Triton-X100 and 1 mM DTT, and then treated with assay buffer
(GME buffer, 16.5 mM KPO4, pH 7.8 2.2 mM ATP, and
1.1 mM DTT). Luciferase activity was measured for 30 seconds
using an AutoLumat luminometer (Berthold Technologies Co.,
Oak Ridge, Tennessee, USA) [10,59].
To assess the effect of betaglycan on the bioactivity of each
mutant, LbT2 cells were transfected with 0.5 mg of rat betaglycan-
c-myc expression plasmid (provided by Dr. Fernando Lopez
Casillas, Howard Hughes Medical Institute, Maryland, USA) or
0.5 mg of empty vector pCDNA3.0 using Lipofectamine 2000
(Invitrogen, Carlsbad, California, USA) in 24-well plates. Betagly-
can transfection efficiency was tested by immunoblot using
monoclonal anti-c-myc antibody. Cells were treated with the
appropriate ligand 24 hours after transfection.
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PLoS ONE | www.plosone.org 12 March 2010 | Volume 5 | Issue 3 | e9457
Immunoprecipitation of Receptor ComplexesCOS7 cells were utilized for immunoprecipitation experiments.
Cells were cultured in DMEM supplemented with 10% fetal
bovine serum and 1% penicillin/streptomycin. At confluency, cells
were split to 26106 cells per 60-cm2 dish. Twenty-four hours later,
cells were transfected with 6 mg of rat ActRIIB-HA expression
plasmid (provided by Dr. Kelly Mayo, Northwestern University,
Evanston, Illinois, USA) and either 6 mg of empty vector pCDNA3
or mouse betaglycan-c-myc expression plasmid. The next day,
normal growth media was added to allow the cells to recover, and
after an additional 24 hours, cells were treated with 100 ng/ml of
the concentrated media. Immunoprecipitation was carried out as
reported previously [10].
Binding AssayPurified human inhibin A or activin A was biotinylated using
the EZ-link Micro Sulfo-NHS-LC-biotinylation Kit (Pierce,
Rockford, Illinois, USA). The binding of biotinylated inhibin A
or activin A to activin type IIB receptor (ActRIIB) was measured
using an enzyme linked immunosorbent assay (ELISA). 50 ng of
ActRIIB in 50 ml of 20 mM HEPES buffer, pH 7.4 was dried
overnight in each well of a 96-well microtiter plate (Nunc
MaxiSorpTM, Fisher Scientific, Pittsburgh, Pennsylvania, USA).
Each well was blocked for 30 min with 100 ml of 0.5% BSA in
PBS, aspirated and washed with 250 ml of PBS. Each well was
then incubated for 1.5 hours with 50 ml of various concentrations
of biotinylated inhibin A in TBST (Tris-buffered saline, pH 7.4,
containing 0.05% Tween 20) at room temperature with shaking.
Wells were aspirated and washed three times with 250 ml of TBS-
T buffer. Each well was then incubated for 30 min with 100 ml of
a 0.1 mg/ml solution in PBS of streptavidin-coupled horseradish
peroxidase (Pierce Chemical Co, Rockford, IL, USA). Wells were
aspirated, washed three times and developed for 15 min with
100 ml of 1-step Ultra TMB (3,39,5,59-tetramethylbenzidine)
obtained from Pierce Chemical Co. (Rockford, IL, USA). Color
development was stopped by the addition of 100 ml 2 M sulfuric
acid. The plate was read using plate reader (Synergy HT Multi-
Mode Microplate Reader, Biotex, Houston, TX, USA) at
450 nm.
Competition Binding AssayThe competition binding of culture media from cells
expressing wild type activin A and activin A/inhibin A chimera
mutants to the ActRIIB was carried out using biotinylated activin
A (EC50 = 1.1 nM based on the biotinylated activin A standard
binding curve shown in Fig. S7A). The competition binding of
culture media from cells expressing wild type inhibin A, chicken
inhibin A, and all of the inhibin a-subunit deletion mutants to the
ActRIIB was carried out using biotinylated inhibin A
(EC50 = 2.73 nM based on the biotinylated inhibin A standard
binding curve shown in Fig. S7B). Different concentrations of
inhibin A in the media were mixed with biotinylated inhibin A in
the binding assay system. The binding assay procedure is the
same as described above. The concentration of the mature
inhibin A or activin A mutants was calculated based on the
immunoblot.
StatisticsValues are reported as the means 6 SD. IC50 values were
determined using the sigmoidal dose-response (variable slope)
curve. Statistical analyses were performed using Prism software
(Version 4.0a, GraphPad Software, San Diego, California, USA).
The F-test was used for the comparison of the IC50 values in each
different group. Statistical significance was reported if p,0.05.
Comparison of the densitometric analysis between different groups
was determined using two-tail unpaired t-test. Statistical signifi-
cance was reported if p,0.05.
Supporting Information
Figure S1 Schematic representation of the human and chicken
inhibin a and human bA pre-pro subunits. i, The entire human
pro-aN-aC is depicted. The full length unprocessed subunit is
366 amino acids. The mature domain (aC) is contained in the 39
end of this subunit and is cleaved by the proconvertase enzyme at
amino acid 232. The red color shows the human aC domain. The
N-terminal extension region of the human a-subunit is shown in
green bar, the proline-rich ‘‘wrist region’’ of the human a-
subunit is shown in blue bar. ii, The entire chicken pro- aN-aC is
depicted. The full length unprocessed subunit is 328 amino acids.
The mature domain (aC) is cleaved by the proconvertase enzyme
at amino acide 215. The yellow color shows the chicken aC. The
N-terminal extension region of the chicken a-subunit is shown in
pink bar. iii, The entire human pro-bA is depicted. The full
length unprocessed subunit is 426 amino acids. The mature
domain (bA) is contained in the 39 end of this subunit and is
cleaved by the proconvertase enzyme at amino acide 310. The
black color shows human bA. The white bar represents the N-
terminal region of the mature bA-subunit, with the crosshatch
pattern bar indicating the wrist a-helical region. The black bars
show the antibody binding sites that are indicated for the R1,
PO14, and PO23 inhibin a-subunit-specific monoclonal anti-
bodies, as well as for pAb bA. The proconvertase enzymeclea-
vage sites were presented by black arrow. The symbol indicates
glycosylation sites.
Found at: doi:10.1371/journal.pone.0009457.s001 (0.34 MB
PDF)
Figure S2 Large-scale estimate of inhibin/activin phylogeny. A)
Maximum likelihood (ML) estimate of phylogeny for genes
retrieved from annotated animal genomes by BLAST (Altschul
et al., 2009) searches. The alignment included only the TGFbdomain of genes that have human inhibin/activin genes as their
top hit (they all satisfied the bidirectional best hit criterion). This
analysis was used the WAG (Whelan and Goldman, 2001) model
with the a proportion of sites assumed to be invariant and the
remaining sites at different rates drawn from a -distribution (with a
shape parameter estimated by ML). Support for clades reflects the
percentage of 100 bootstrap replicates. B) Alignment used for part
A of this figure, with the 3 amino acid insertion that unites the
lancelet (Branciostoma) a-subunit-like protein with the vertebrate
(in this alignment, human and zebrafish) inhibin a-subunit
highlighted using yellow. C) Alignment for the lancelet a-
subunit-like protein with the lancelet b protein mature domain
sequence. D) Alignment for the mature domain sequences between
the human inhibin a-subunit, chicken inhibin a-subunit and
human inhibin bA subunit. The red box that used in the C) and D)
highlighted the regions that we focused on.
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PDF)
Figure S3 Detailed version of the inhibin/activin phylogeny
presented in Fig. 1C. The ML analysis used the JTT+ f+invariant
sites model, as described in the text. Support for specific groups
reflects the percentage of 100 bootstrap replicates (only values
$50%). The version in the text indicates the occurence of indel
mutations, including the indels shown to have a major functional
impact.
Evolution of Inhibin a-Subunit
PLoS ONE | www.plosone.org 13 March 2010 | Volume 5 | Issue 3 | e9457
Found at: doi:10.1371/journal.pone.0009457.s003 (0.08 MB
PDF)
Figure S4 Detail of mutations within human inhibin aC and
bA-subunits. A) The schematic representation of wild-type human
aC and human bA-subunits. The letters ‘‘A’’, ‘‘B’’. ‘‘C’’, ‘‘D’’, ‘‘E’’
and ‘‘F’’ denote the six candidate regions targeted for deletion. B)
The residue numbers and sequences for the six regions targeted for
deletion and used for generating the inhibin a-subunit deletion
mutants and inhibin bA-subunit chimera mutants. C) Design and
naming conventions for the a-subunit and b-subunit mutants.
Found at: doi:10.1371/journal.pone.0009457.s004 (0.32 MB
PDF)
Figure S5 Negative control for the Fig. 3B and Tandem MS-MS
fragment mass mapping. A) Left, immunoblot of media from cells
expressing wild type human a-subunit alone and human inhibin A
under non-reducing or reducing conditions. Middle, immunoblot
of media from cells expressing human free a or human inhibin A
that were labeled with [35S]-cysteine for 48 hours. Media were
collected and immunoprecipitated with an anti-a-subunit mono-
clonal antibody PO23 and then subjected to SDS-PAGE under
non-reducing conditions and autoradiography. Right, negative
control immunoblot of mouse IgG-immunoprecipitated [35S]-
cysteine labeled media. Labels 1, 2, 3 and 4 indicate media from
cells expressing aHwt/aHwt, aHwt/bA, aChwt/aChwt and
aChwt, respectively. B) The culture media from the chicken a-
subunit alone expression cell was collected and immunoprecipi-
tated with an anti-a-subunit monoclonal antibody PO23 and
subjected to SDS-PAGE under non-reducing conditions. The gel
was stained with Coomasie. The band marked with ‘‘*’’ is
supposed chicken a-a homodimer band The molecular weight is
around 30 kDa. C) The band was digested with MS grade trypsin
and subjected to tandem MS- MS analysis. Using the SwissProt
database, the resulting fragment from the band mapped to the
chicken inhibin a-subunit mature domain. The matched fragment
is highlighted in red.
Found at: doi:10.1371/journal.pone.0009457.s005 (3.97 MB
PDF)
Figure S6 Immunoblots of media from cells expressing wild type
human and chicken inhibin A and various human a-subunit
deletion mutants run under reducing conditions. Labels 19, 29, 39,
49 and 59 indicate media from cells expressing aHwt/bA, aChwt/
bA, aHext-/bA, aHPWR-/bA and aHext-PWR-/bA, respective-
ly. A) Blot was detected by an anti-a-subunit monoclonal antibody
PO23. B) Blot was detected by anti-bA-subunit polyclonal
antibody.
Found at: doi:10.1371/journal.pone.0009457.s006 (1.58 MB
PDF)
Figure S7 Standard curves for biotinylated activin A/inhibin A
binding to ActRIIB. A) Standard curve for biotinylated activin A;
EC50 = 1.10 nM. Related to Fig. 4A. B) Standard curve for
biotinylated inhibin A; EC = 2.73 nM. Related to Fig. 4C.
Found at: doi:10.1371/journal.pone.0009457.s007 (0.12 MB
PDF)
Figure S8 Negative and positive controls for the co-immunopre-
cipitation experiments presented in Fig. 4B. COS7 cells were
transiently transfected with ActRIIB-HA with either pCDNA3 or
betaglycan-c-myc (BG-c-myc). A) Cells were treated with 100 ng/ml
human activin A or empty vector culture media (EV media). The cell
lysate was immunoprecipitated by monoclonal anti-HA or mouse
IgG, followed by immunoblot with anti-bA-subunit polyclonal
antibody. B) Cells were treated with 100 ng/ml human activin A
(panel 1); empty vector (panel 2), human wild type inhibin A (panel 3);
chicken wild type inhibin A (panel 4); or culture media from cells
expressing human inhibin A deletion mutants aHext-/bA (panel 5),
aHPWR-/bA (panel 6), or aHext-PWR-/bA (panel 7). Proteins were
detected in the cell lysates by immunoblotting with either monoclonal
anti-c-myc or monoclonal anti-HA antibodies.
Found at: doi:10.1371/journal.pone.0009457.s008 (1.56 MB
PDF)
Table S1 Species database information for TGFb family
members analyzed in Fig. 1A and Fig. 1B.
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PDF)
Table S2 Species database information for inhibin a-subunit
analysis in Fig. 1A and Fig. 1C.
Found at: doi:10.1371/journal.pone.0009457.s010 (0.07 MB
PDF)
Table S3 Oligonucleotide primers used for deletion mutagenesis.
Found at: doi:10.1371/journal.pone.0009457.s011 (0.04 MB
PDF)
Acknowledgments
We would like to thank Candace Tingen and Min Xu for their helpful
discussions.
Author Contributions
Conceived and designed the experiments: JZ ELB LJGJ TJ TW.
Performed the experiments: JZ SK MA EYX RWC SJL BCM. Analyzed
the data: JZ ELB SK TW. Contributed reagents/materials/analysis tools:
JZ TW. Wrote the paper: JZ ELB TW.
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